FULL PAPER
Extracellular Vesicles
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Tangential Flow Microfluidics for the Capture and Release
of Nanoparticles and Extracellular Vesicles on Conventional
and Ultrathin Membranes
Mehdi Dehghani, Kilean Lucas, Jonathan Flax, James McGrath,* and Thomas Gaborski*
Membranes have been used extensively for the purification and separation of biological species. A persistent challenge is the purification of
species from concentrated feed solutions such as extracellular vesicles
(EVs) from biological fluids. Investigated is a new method to isolate microand nanoscale species termed tangential flow for analyte capture (TFAC),
which is an extension of traditional tangential flow filtration. Initially,
EV purification from plasma on ultrathin nanomembranes is compared
between both normal flow filtration (NFF) and TFAC. NFF results in rapid
formation of a protein cake which completely obscures any captured EVs
and also prevents further transport across the membrane. On the other
hand, TFAC shows capture of CD63 positive small EVs with minimal
contamination. The use of TFAC to capture target species over membrane
pores, wash, and then release in a physical process that does not rely
upon affinity or chemical interactions is explored. This process is studied
with model particles on both ultrathin and conventional thickness membranes. Successful capture and release of model particles is observed
using both membranes. Ultrathin nanomembranes show higher efficiency
of capture and release with significantly lower pressures indicating that
ultrathin nanomembranes are well-suited for TFAC of delicate nanoscale
particles such as EVs.
M. Dehghani, Dr. T. Gaborski
Department of Microsystems Engineering
Rochester Institute of Technology
Rochester, NY 14623, USA
E-mail: trgbme@rit.edu
M. Dehghani, Dr. T. Gaborski
Department of Biomedical Engineering
Rochester Institute of Technology
Rochester, NY 14623, USA
K. Lucas, Dr. J. McGrath, Dr. T. Gaborski
Department of Biomedical Engineering
University of Rochester
Rochester, NY 14627, USA
E-mail: jmcgrath@bme.rochester.edu
Dr. J. Flax
Department of Urology
University of Rochester Medical School
Rochester, NY 14642, USA
The ORCID identification number(s) for the author(s) of this article
can be found under https://doi.org/10.1002/admt.201900539.
DOI: 10.1002/admt.201900539
Adv. Mater. Technol. 2019, 1900539
1. Introduction
Tangential flow filtration (TFF) is used
extensively in bioprocessing.[1] In this
method, a feed solution containing a species of interest flows tangentially over
a selective membrane with some fraction of the flow also passing through the
membrane. If the species of interest is to
be retained behind the membrane, TFF
can be used to remove impurities or to
concentrate the species in the feed solution.[2,3] If the volume lost through transmembrane flow is resupplied to the feed
channel as fresh buffer (diafiltration),
TFF can be used for buffer exchange.[4,5]
TFF can also be used to partially purify
a species that emerges in the filtrate,
although the product typically requires
final purification by column or membrane
chromatography.[6–10]
The advantage of TFF over normal
flow filtration (NFF) is that the tangential
flow component disrupts the formation
of a concentration polarization layer that
builds as species are rejected by the membrane.[11] Without a tangential component,
this polarization layer will eventually form
a “cake” layer on the membrane with its own separation properties and significantly reduced permeate flux.[12] With TFF filtration however, it is possible to identify conditions for which both
the flux and transmembrane pressure (TMP) are steady with
time.[13] Under these conditions, filtration can, in principle,
continue indefinitely.
Our laboratories develop ultrathin porous membranes for
a range of applications including separations.[14–17] Ultrathin
membranes are best defined as materials with pores on the
same order as, or larger than, the membrane thickness.[18]
These have been made with a variety of materials including
silicon, silicon-nitride, silicon dioxide, graphene, and grapheneoxide.[19–24] We have recently demonstrated that the high permeability of ultrathin membranes causes them to foul rapidly in
NFF, with initial pore blockage events quickly followed by cake
filtration.[25] We showed the same fouling phenomena occurs
with both particle and protein solutes when used in NFF.[25,26]
To extend the capacity of ultrathin membranes in separations, we have recently examined their performance in TFF.
Working with undiluted serum and nanoporous silicon nitride
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(NPN) membranes, we made the surprising discovery, reported
here for the first time, that 60–100 nm extracellular vesicles
(EVs) are captured in the pores of ultrathin membranes with
little evidence of protein fouling.[27] Our discovery inspired
a closer look at the mechanisms and potential utility of capturing nanoparticle-sized analytes from biofluids in the pores
of ultrathin membranes.
EVs are secreted from tissue cells into all body fluids, and EVs
that are <100 nm are typically, but not exclusively, exosomes.
Exosomes contain the largest pool of extracellular RNA (exRNA)
in biofluids, and are thus valued both for their diagnostic and
therapeutic potential.[28–33] The conventional method for exosome
purification is ultracentrifugation, although many alternative
strategies have been proposed, including TFF.[34–37] Out of respect
for the careful criterion used to define exosomes, we will refer to
<100 nm EVs as small EVs (sEVs) rather than exosomes.[38]
We propose a novel method for the extraction of nanoparticle
species from biofluids which we call tangential flow for analyte capture (TFAC). In this method, sEVs and similarly sized
analytes are captured in the pores of an ultrathin membrane
where they can be washed and released with additional flows.
TFAC resembles bind/elute purification strategies although it
distinguishes itself from affinity chromatography because the
binding is purely physical. TFAC does not require engineered
surface chemistries for capture or chemical treatments for elution. The purpose of the current report is to demonstrate the
basic principles of TFAC using model particles. We also test
the hypothesis that ultrathin membranes are ideally suited for
TFAC because they facilitate capture and release at lower pressures than conventional thick membranes.
2. Results
2.1. Tangential Flow for Particle Capture
The system and scheme for particle capture and release is
shown in Figure 1. As in our prior work,[39–42] we used layerby-layer assembly (Figure 1A) to construct microfluidic devices
(Figure 1B) with membranes separating top and bottom flow
channels. The only difference is that we used a clamped system
for both polycarbonate track-etch (PCTE) systems and NPN systems instead of a fully bonded devices. This enables the removal
and inspection of PCTE membranes or NPN chips by scanning
electron microscope (SEM) after use. Particle capture (Figure 1C)
was performed using two syringe pumps: a positive pressure
pump providing a constant sample supply flow rate into the
input channel of the device, and a negative pressure pump at
the output channel exit side controlling a smaller, steady rate of
ultrafiltration through the membrane. The difference between
the supply and ultrafiltration rates exited the top channel as waste
and provided the tangential flow needed to prevent fouling.[11,13]
The inlet port on the bottom channel was blocked for all experiments. After capture, nonadsorbed contaminants could be cleared
by replacing the sample with a rinse buffer while maintaining
the TMP (Figure 1D) and captured analytes could be released by
operating both pumps under positive pressure (Figure 1E).
2.2. sEV Capture from Undiluted Serum
The initial discovery of analyte capture occurred with experiments
on undiluted serum. In these experiments, we showed that the
Figure 1. TFAC technique for isolation of particles. A) Microfluidic devices are assembled through a layer stack process, in which channels and other
featured are patterned into PDMS sheets. B) These layers are then formed into the device through thermal bonding or stacking and clamping. C) The
sample is passed across the surface of the membrane and a TMP generated by syringe pumps drives particle motion toward the membrane. Contaminating particles pass through pores or are swept downstream while the particles are retained on the membrane surface. D) The cleaning buffer is then
passed through the input channel under the same flow condition as the capturing step to wash the channel and membrane surfaces of any remaining
contaminants. E) The TMP is then reversed, releasing the particles from the membrane where they are then swept downstream and collected.
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Figure 2. sEVs captured from undiluted blood plasma. A) SEM images showing the thinness and high porosity of NPN. B) In NFF, a protein cake
of ≈8 µm rapidly builds up on the membrane surface. C) After capturing and cleaning steps of TFAC, small vesicles are captured on the membrane
surface with minimum fouling. D) Nanogold-conjugated anti-CD63 antibody labels an EV captured in a pore multiple times, indicating it is likely a
CD63 positive sEV. Note: the fragmented appearance of the surface results from the use of a limited amount of gold (3 nm) to avoid obscuring the
gold label on the antibody (18 nm). By contrast 10 nm of gold was sputtered on the samples to avoid charging effects in SEM in both (B) and (C).
Scale bar = 200 nm for (A)–(C). Scale bar = 50 nm for (D).
filtration of undiluted serum is difficult in NFF (Figure 2B),
causing an 8 µm cake of serum protein and salts to foul the membrane and allowing the passage of only 10 µL of a 1 mL sample.
However, upon passing the undiluted serum across the
membrane in tangential flow, we observed a significant reduction in the protein build-up on the membrane, showing captured particles (Figure 2C). Human plasma and serum contain
different types of particles including EVs and lipoproteins. Lipoproteins and EVs cannot be distinguished only by their physical
properties since their size and density closely overlap.[43,44]
However, further analysis of the captured particles with immunostaining showed that some of the particles were positive
for CD63 which is a common sEV surface protein and is not
expressed on lipoproteins. We did not attempt rinse or release
steps with undiluted serum, instead we turned to the following
experiments with model systems to confirm and study the capture phenomena under defined conditions.
2.3. Microporous Track-Etch Capture of Fluorescent Particles
We first explored the particle capture phenomena at the microscale using microporous polycarbonate track-etched (mPCTE)
membranes with 8 µm pores and 10 µm particles. At this scale,
we were able to image individual particle capture events in
fluorescence microscopy (Figure 3). Before flow (T0; Figure 3B)
there were no particles on the membrane. With a steady supply
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rate of 90 µL min−1 and ultrafiltration rate of 10 µL min−1, particles began to accumulate on the membrane, primarily drawn
directly to the pores (see electron micrograph in Figure 3C,
bottom panel), and the fluorescence steadily increased over
time (Figure 3C). The capturing process was then stopped at
T1 resulting in immediate release of particles loosely held on the
membrane and a distinct, sudden drop in fluorescence (light
blue line Figure 3A). Finally, the flow was reversed by switching
the ultrafiltration pump to infusion mode, resulting in a directional shift for the bottom flow. The bottom flow rate was then
increased to provide a high TMP in an attempt to fully release
the remaining particles, although a fraction remained irreversibly bound resulting in a residual fluorescence after the experiment T2 (Figure 3D). Electron microscopy (Figure 3D, bottom
panel) shows that most of these particles were not associated
with pores and thus were nonspecifically adhered to the surface
of the membrane through surface interactions. More than 90%
of the particles captured were released (Figure S2 and Movie S2,
Supporting Information), suggesting this method has promise
for the purification of microscale particles.
2.4. Nanoporous Track-Etch Capture of Fluorescent Nanoparticles
Having demonstrated capture using modified tangential flow
in a microscale system, experiments were performed to show
capture and release at the nanoscale. Track-etch membranes
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Figure 3. Microscale experiments with 10 µm fluorescent particles and 8 µm pore size polycarbonate track-etch (mPCTE) membranes. A) Fluorescence
intensity/number of particles–time plot showing an increase in the intensity signal/number of particles during the capturing step and a decrease for
releasing step. B–D) Before capturing, after capturing, and after releasing panels, respectively, including schematic images (top), fluorescent images
(middle), and SEM images (bottom). The red diagonal in fluorescent panel (C) shows releasing of captured particles by pausing the pump to change
the flow configuration. Particles in panel (C) are labeled with either single or double arrowheads, indicating nonspecifically bound and captured particles, respectively.
with 80 nm pores (nPCTE) were used to capture 100 nm fluorescent nanoparticles. Because of the significant increase
in membrane resistance compared to mPCTE, flow rates of
5 µL min−1 (sample supply) and 2 µL min−1 (ultrafiltration)
were now used for capture. This was followed by washing with
clean buffer to remove any nonspecifically bound particles
before the releasing in a backwash step (5 µL min−1 backflow).
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During the capture phase of the experiments, the fluorescence intensity curves displayed similar behavior to the microscale experiments, with a steady increase throughout the capture
period (Figure 4A). Unlike the mPCTE experiments however,
there was no observable loss of fluorescence after the release of
TMP at the end of the capture phase. A fraction of loosely associated particles, either on the surface or in suspension above
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Figure 4. Nanoscale experiments with 100 nm fluorescent particles and 80 nm median pore size polycarbonate track-etch (nPCTE) membranes.
A) Fluorescent intensity analysis (solid line) showing the gradual increasing and decreasing in the fluorescent signal during the capturing step and
cleaning step, respectively, followed by a sharp drop as nanoparticles were released (the dash line shows the intensity change during the experiment
in the absence of the TMP). Scale bar on fluorescence image insets = 50 µm. B–D) Electron micrographs showing before capturing, after capturingcleaning, and after releasing panels, respectively.
the surface (Figure 4C), were removed with a wash step. Flow
reversal did not fully remove all the particles captured on the
membrane as some were lodged deep within pores (Figure 4D
and Figure S3, Supporting Information), but the system did
return to within ≈85% of the baseline fluorescence value.
In order to assess the role of the applied TMP on capturing,
experiments were performed in the absence of active TMP
(dashed line, Figure 4A). To achieve this, supply flow was performed as before, but the ultrafiltration pump was not used to
generate active transmembrane flow. While the change in fluorescence intensity showed an increase in particles, the maximum
measured intensity was only 50% of the system with active TMP,
which indicates that TMP is the driving force of particle capture.
2.5. Nanoporous Silicon Nanomembrane Capture
of Fluorescent Nanoparticles
Our original observations of EV capture from serum (Figure 2)
were obtained with 100 nm thick NPN membranes.[27] It is
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important to note that PCTE membranes used are approximately 60 times thicker compared to ultrathin nanomembranes. Thus, our next set of studies replicated the experimental
conditions used with nPCTE on NPN (5 µL min−1 supply;
2 µL min−1 ultrafiltration) with similar pore sizes (80 nm
median) and total number of pores actively filtering materials were of the same order (nPCTE = 4 × 107 pores mm−2;
NPN = 9.2 × 107 pores mm−2), which resulted in a slightly larger
membrane area for the nPCTE membranes (4 mm2) compared
to the NPN membranes (1.4 mm2). Therefore, membrane thickness and membrane surface chemistry are the key parametric
differences between experiments on nPCTE versus NPN.
The capture and release intensity curves (Figure 5A) with
NPN show similar trends to nPCTE with some interesting
differences. There is again an increase in fluorescence intensity on the membrane during the capture phase followed by
a sudden loss of particles when the flows are stopped. After a
rinse with clean buffer, the intensity returns to within ≈95% of
the baseline, which is slightly better than that seen with nPCTE
(Figure 5A, inset). A control in the absence of TMP (Figure 5A,
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Figure 5. Nanoscale experiments with 100 nm diameter fluorescent particles and 80 nm median pore size NPN membranes. A) Fluorescent intensity
analysis (solid line) showing the increasing during the capturing step, and then decreasing in the cleaning step following by a drop as nanoparticles
were released (the dash line showing the intensity changes during the experiment in the absence of the TMP). Scale bar on fluorescence image
insets = 50 µm. B–D) Electron micrographs showing before capturing, after capturing-cleaning, and after releasing panels, respectively.
dashed line) showed once more that the capture process is
driven by TMP.
Electron microscopy was again performed to better understand the capture process. The membrane showed high pore
density (Figure 5B), in contrast with track-etch membranes
(Figure 4B), and a distribution of pore sizes with median of
80 nm (Figure S4, Supporting Information). As expected, the
majority of the 100 nm particles captured remained on top
of the pores (Figure 5C). A small proportion of particles persisted on the membrane after the releasing step, and these all
appeared to be captured within pores (Figure 5D).
In order to estimate particle concentrations throughout the
capture and release process, calibration curves for both NPN
and nPCTE experiments were made by correlating the fluorescent intensity to the number of particles on the membrane
(Figure S5, Supporting Information). These curves allowed for
the direct comparison of membrane performance for particle
capture and release (Table 1). We estimate that track-etch membranes capture ≈2.6 × 106 particles from an available population
of 5 × 107 and released 60% of the particles captured. By contrast, silicon nanomembranes captured ≈8.6 × 106 particles from
the same solution and released 68% of the captured population.
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2.6. Pressure Modeling of Track-Etch and Ultrathin
Silicon Nitride Nanomembranes
We explored the effect of the membrane thickness on TMP in
our studies both analytically and experimentally. Experiments
were conducted with nanoporous track-etch membranes and
ultrathin NPN nanomembranes. Pressure sensors were placed
upstream and downstream on either side of the membrane
(Figure 6A) and the pressures were monitored under flow
conditions equivalent to the capture experiments (5 µL min−1
supply, 2 µL min−1 ultrafiltration). Results for both nPCTE and
NPN compared favorably to predictions of the Dagan equation—a modified Hagen–Poiseuille equation that also applies
Table 1. Nanoporous track-etch (nPCTE) versus silicon nanomembrane
(NPN) captured and released particle counts.
Membrane
Captured particles
Released particles
TE membranes
2.52 +/− 0.24 × 106
1.56 +/− 0.43 × 106
NPN membranes
8.62 +/− 3.15 × 106
5.89 +/− 1.98 × 106
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Figure 6. Theoretical and experimental pressure drops across nanoporous polycarbonate track-etch membranes (nPCTE) and NPN membranes. A) Diagram of the pressure monitoring system showing the position of the pressure sensors and the direction of flow. All flow was performed at 10 µL min−1
through the membrane with a syringe pump pushing on the top channel and a syringe pump pulling on the bottom channel. The pressure sensors
were positioned 5 cm above and below the membrane. B) Comparison of pressure drops across the track-etch and NPN membranes. Blue = Dagan
predicted, homogenous distribution pressure drop. Red = experimental data. Logarithmic scale used for comparison. C) COMSOL model of pressure
in a track-etch system showing a large pressure drop across the membrane. D) COMSOL model of pressure in an NPN system showing almost no
pressure drop across the membrane, in stark contrast to the track etch system. COMSOL simulations were performed using the Free and Porous Flow
toolbox with a Darcy’s permeability calculated for this system.
to ultrathin membranes.[14,40,45] The Dagan equation gives the
pore resistance as
R pore =
µ
8 L 
3 +   
π  r 
r 3 
(1)
where μ is the fluid viscosity [Pa s−1], r is the pore radius [m],
and L is the pore length [m]. The total membrane resistance R
is calculated by adding the resistance for each pore in the membrane in parallel (9.2 × 107 pores for NPN and 4 × 107 pores
for nPCTE) and the anticipated pressure drop is then found by
multiplying by the flow rate
∆P = Q ⋅ R
(2)
The comparison of this estimate with experimental results
(Figure 6B) showed that a simple analytical approximation is
sufficient for predicting the TMP drop that could be experienced in the system. These results were compared to an analytical model of pressure drop (Figure 6B) as well as COMSOL
Multiphysics simulations (Figure 6C,D) to illustrate the pressure gradients and streamlines in the system.
3. Discussion
In this work, we introduced a new method for sample purification in which particles are captured on the surface of a
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membrane in tangential flow, washed to remove contaminants,
and then released in a controlled fashion where they can be further analyzed, concentrated, or processed.
We call this process TFAC and while the process resembles bind and elute strategies found in column or membrane
chromatography, it relies on physical interaction, rather than
chemical affinity, for capture. Similarly, TFAC requires physical
release through back-flow for elution, rather than chemical
treatments to disassociate chemical bonds formed during capture. As the release of chemical bonds in affinity schemes can
often be destructive and incomplete, there are clear advantages
for physical capture and release.
Proof-of-concept experiments using fluorescent particles
on both PCTE and NPN membranes showed successful capture and release of particles. We have shown that NPN membranes outperform PCTE membranes for capture and release
with poly­styrene nanoparticles. Our analytical and experimental
comparison showed that the greater thickness of PCTE compared to NPN caused higher TMP. This high pressure drives
nanoparticles into the membrane bulk where they disappear
from view and are more difficult to recover (Figure 4).
One potential application of TFAC utilizing ultrathin membranes as a microfluidic-based technique would be isolation
of EVs. Studies have indicated that not only the RNA content
of these vesicles but also their protein varies by cell of origin
as well as by the pathologic state of these cells.[46–50] This differential cell state specific and cellular origin-based content
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Figure 7. TFAC Illustration showing capturing, cleaning, and releasing steps.
indicates that EVs could serve as biomarkers of disease. These
biomarkers could be used for a range of clinical purposes
including disease screening, predicting a priori disease responsiveness to treatment, and monitoring response to treatment.
Currently, the “gold standard” method for isolating EVs from
biofluids is ultracentrifugation, which requires large volumes
of biofluid (>25 mL), long processing times, expensive instrumentation, and trained technicians. Gel precipitation and size
exclusion chromatography have been developed that remove
the need for ultracentrifugation and allow EV isolation in a
benchtop centrifuge, but these methods suffer from low yield
and/or contamination with co-precipitated proteins.[51–55] The
high protein contamination from these methods prevents the
use of EV proteins as biomarkers in addition to RNA. The result
from our plasma isolation experiment by ultrathin nanomembranes showing capture of EVs with minimal contamination
suggests promising potential of TFAC for isolation of EVs with
high purity (Figure 7).
Additionally, the pore size of the ultrathin NPN membranes
can be tuned to capture different subpopulations of EVs that
vary in size.[56,57] This includes microvesicles, exosomes, and
apoptotic bodies which are diagnostically informative.[58] In all
cases, TFAC method eliminates the necessity of preprocessing
biofluids which can be both time-consuming, results in sample
loss, and often requires specialized equipment reducing the
utility of these particles in point of care diagnostic devices.[59,60]
Another potential application of TFAC would be a membrane-based “in situ” analysis to detect EVs carrying cancer biomarkers among a larger population using the same membrane
for capture, labeling, and imaging by fluorescence microscopy.
TFAC using NPN membranes showed that captured particles
were associated with membrane surface, rather than trapped
in a bulk-matrix which means that the captured particles can
be analyzed directly on the membrane. Furthermore, TFAC
captured EVs from whole plasma with minimal contamination
(Figure 2C) as opposed to rapidly formed cake on the membrane
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by NFF which increases the sensitivity and specificity of EVs
biomarker detection. Also, the excellent optical properties of
ultrathin inorganic membranes like NPN, would also be key to
enabling this application.[61] In comparison, track-etch membranes lack this optical transparency and as the current study
indicates, trap EVs below the membrane surface, together precluding the ability to detect specific diagnostic markers directly
in and on the EVs captured on the membranes.[62]
The abundant presence of EVs and lipoproteins with similar physical characteristics such as size and density in blood
makes it one of the most difficult body fluids to isolate EVs
from.[63] Co-purification of EVs and lipoproteins has been
observed using other size-based separation methods such as
size exclusion chromatography and density gradient ultracentrifugation.[43,44] Therefore, contamination of EVs with blood
lipoproteins may occur using TFAC. However, affinity-based
separation can be performed downstream to decrease the blood
lipoproteins contaminants level when EV samples with high
purity is desired.[64,65] On the other hand, lipoproteins are not
likely to be present in cell-conditioned media, especially when
cells are cultured with serum-free media.
4. Conclusion
In this work, we have developed a method called TFAC to
capture and release of particles. We contend that ultrathin
membranes are ideally suited for TFAC for two reasons: 1)
operating pressures are orders-of-magnitude lower for ultrathin
membranes than for membranes with conventional thicknesses (1–10 µm) and 2) captured particles are associated with
a surface, rather than trapped in a bulk-matrix and 3) higher
efficiency of capture and release of particles. Experiments
performed in NFF with human plasma demonstrated formation of a protein cake on the surface of ultrathin membranes.
However, testing human plasma in TFAC mode resulted in
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capturing EVs with minimal contamination. Captured vesicles
were further labeled in situ, providing a convenient platform
for downstream detection and analysis. Together, these findings
suggest promising potential of TFAC for both isolation of EVs
and biomarker detection on captured EVs.
5. Experimental Section
Fabrication of NPN Membranes: The fabrication steps for NPN
nanomembranes have been published previously.[19] Briefly, a silicon
wafer was coated with a three-layer stack of silicon nitride (SiN),
amorphous silicon, and silicon dioxide. A porous nanocrystalline silicon
(pnc-Si) layer was formed on top of SiN via rapid thermal annealing. The
nanopores present in the pnc-Si were transferred into the SiN layer by
reactive ion etching. In order to create the free-standing membranes, the
back side of the silicon wafer was etched to the silicon nitride layer using
ethylene diamine pyrocatechol.
NPN Device Fabrication: Polydimethylsiloxane (PDMS) sheets
(Trelleborg Sealing Solutions Americas, Fort Wayne, IN) were used to
create microfluidic devices. Custom ordered 100 µm and 300 µm thick
restricted grade sheets were patterned using a Silhouette Cameo digital
craft cutter (Silhouette America, Oren, UT).[66] The patterned silicone
sheets were assembled into layer stack devices by aligning the patterned
layers (Figure 1A and Movie S1, Supporting Information). NPN
membrane chips (300 µm thick) were sandwiched between stacked
layers and the final device was clamped to seal it for flow.
PCTE Device Fabrication: As a representative of conventional thickness
membranes, commercial PCTE membranes with pore sizes of 8 µm and
80 nm were utilized (Sterlitech, WA, USA). In order to have a sealed
system for track-etch membranes, the above-described microfluidic
device was modified. Holes were drilled in polycarbonate slabs for
accessing the bottom channel of the device, while the PDMS slabs were
punched for flowing to the top channel. 100 and 300 µm thick patterned
PDMS sheets were used for bottom and top channels, respectively. In
order to prevent leaking in the system, the PCTE membranes covering
the entire device were sandwiched between the top and bottom layers
using a clamp (Figure S1, Supporting Information).
sEV Capture from Plasma: NFF: sEV experiments were performed
using purified human plasma (Equitech-Bio, Inc., Kerrville, TX). NFF
experiments were performed using NPN chips with 50 nm thick freestanding membranes, with an average pore diameter of 50 nm and a
porosity of 15% in a SepCon centrifuge cup (SiMPore Inc., Rochester, NY).
A 500 µL sample of undiluted plasma was spun at 1500 × g through the
membrane and the chip was extracted from the device. The chip was
allowed to dry and was then imaged by SEM as described below.
TFAC: NPN microfluidic devices were fabricated as described above.
The NPN chip used had a 50 nm thick free-standing membrane with
a 50 nm average pore diameter and a 15% porosity. 1 mL of plasma
was passed tangential to the membrane surface at a rate of 10 µL min−1
using a syringe pump (Chemyx Fusion 200, Chemyx Inc., Stafford,
TX), while fluid was actively pulled through the membrane at a rate
of 2 µL min−1. After processing the full 1 mL volume, the device was
unclamped and the chip extracted. Captured sEVs were labeled for CD63
(Abcam, Cambridge, MA) and imaged via SEM as outlined below.
Capture and Release: Microscale Experiments: Flow experiments were
performed using two Chemyx Fusion 200 syringe pumps (Chemyx
Inc., Stafford, TX). Micron scale experiments with 10 µm polystyrene
green fluorescent particles (Thermo Scientific, USA) were conducted
on 8 µm track-etch membranes. Capturing step was performed using
a sample supply flow rate of 90 µL min−1 and an ultrafiltration/pulling
rate of 10 µL min−1. Captured particles were released by reversed flow of
10 µL min−1 through the membrane.
Nanoscale Experiments: These experiments were conducted using
100 nm polystyrene green fluorescent particles (Thermo Scientific,
USA) on PCTE or NPN membranes with 80 nm median pore size.
Nanoparticles were captured by supply flow rate of 5 µL min−1 and the
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ultrafiltration/pulling flow rate of 2 µL min−1. Input channel was then
cleaned by rinsing buffer to wash away the floating particles under the
same flow condition as the capturing step. Finally, captured particles
were released by reversed flow of 2 µL min−1 through the membranes.
Time-Lapse Video Microscopy: Devices were illuminated with metal
halide lamp source (LE6000 Leica) through differential interference
contrast (DIC) and fluorescein isothiocyanate (FITC) (488 nm Ex/525 nm
Em) filter sets on a Leica DM16000 microscope (Leica Microsystems,
Buffalo Grove, IL) using the 10X objective. Images were collected using
MetaMorph software with a Rolera em- camera (QImaging, Surrey,
BC Canada) for 50 ms exposure time for FITC and 10 ms for DIC. The
measuring and merging channel tool in NIH ImageJ were used for
quantifying the average intensity values and making videos by merging
DIC with FITC images, respectively. Images were taken every minute for
nanoscale experiments and every second for microscale experiments.
Electron Microscopy: After the completion of experiments, the PCTE
and NPN membranes were imaged via electron microscopy. Samples
were prepared for electron microscopy by first removing the membranes
from the device and then allowing them to air dry. Samples were then
mounted and sputter coated with ≈3–10 nm of gold. Scanning electron
micrographs were taken at an accelerating voltage of 10 kV using either
a Hitachi S-4000 SEM or a Zeiss AURIGA SEM.
Supporting Information
Supporting Information is available from the Wiley Online Library or
from the author.
Acknowledgements
M.D. and K.L. contributed equally to this work. The authors acknowledge
Brad Kwarta for the EV capture and release illustration. Research
reported in this publication was supported in part by the National
Science Foundation (IIP 1660177) to J.L.M and T.R.G., Department of
Defense (CA170373) to J.L.M., and the National Institutes of Health
(R35GM119623) to T.R.G.
Conflict of Interest
The authors declare the following competing financial interest: J.L.M.
and T.R.G. are co-founders of SiMPore and hold an equity interest in
this early-stage company commercializing ultrathin silicon-based
technologies.
Keywords
exosomes, extracellular vesicles, nanomembrane, normal flow filtration,
tangential flow for analyte capture, track-etch membranes
Received: June 27, 2019
Revised: August 8, 2019
Published online:
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